Conversions of Formaldehyde-modified 2′-Deoxyadenosine 5′-Monophosphate in Conditions Modeling Formalin-fixed Tissue Dehydration

Abstract

Formalin-fixed, paraffin-embedded specimens typically provide molecular biologists with low yields of extractable nucleic acids that exhibit extensive strand cleavage and covalent modification of nucleic acid bases. This study supports the idea that these deleterious effects are promoted by the first step in formalin-fixed tissue processing—i.e., tissue dehydration with a graded series of alcohols. We analyzed the conversions of formaldehyde-modified 2′-deoxyadenosine 5′-monophosphate (dAMP) by reverse-phase ion-pair, high-performance liquid chromatography and found that dehydration does not stabilize N-methylol groups in the modified nucleotide. Furthermore, spontaneous demodification in a dry state or in anhydrous ethanol can be as fast as it is in aqueous solutions if the preparation is contaminated with salts of orthophosphoric acid. In ethanol, orthophosphates also catalyze formation of abundant N⁶-ethoxymethyl-dAMP, as well as cross-linking and depurination of nucleotides present in the mixture. Identification of the products was performed using ultraviolet absorbance spectroscopy and electrospray ionization Fourier-transform ion cyclotron resonance mass spectrometry. Alternatives to the traditional processing of formalin-fixed tissues are discussed.

Keywords

formalin fixation dAMP mixed acetals crosslinking depurination HPLC ESI FTICR MS

For many decades, the fixation of tissue specimens in 10% formalin (3.7% formaldehyde) has served as the preeminent procedure for tissue morphology preservation in clinics and laboratories. Unfortunately, formalin fixation complicates the characterization of the specimens by modern molecular biological techniques used to reinforce and extend traditional histology. In particular, formalin-fixed, paraffin-embedded tissues typically provide low yields of extractable DNA and RNA that exhibit significant degradation (Goelz et al. 1985; Dubeau et al. 1986; Rupp and Locker 1988) and result in the detection of artificial mutations (Wong et al. 1998; Williams et al. 1999). The chemical bases of these deleterious effects, and ways to prevent or reverse them, have yet to be investigated. This is because, in aqueous solutions, formaldehyde did not appear to cause strand cleavage in either RNA (Haselkorn and Doty 1961; Axelrod et al. 1969) or DNA (Tokuda et al. 1990; Noguchi et al. 1997). Moreover, the primary products of formaldehyde reactions with nucleic acid heterocyclic bases, N-methylol (or N-hydroxymethyl) groups, were shown to fully decompose after a short, high-temperature treatment at neutral pH (Haselkorn and Doty 1961; McGhee and von Hippel 1975). In addition, these methylol groups were found to be fairly inactive in the formation of intra- and intermolecular cross-links (Feldman 1967, 1973) that could be responsible for comparatively stable modifications and for nucleic acid covalent entrapment in formalin-fixed specimens. For example, Solomon and Varshavsky (1985) demonstrated that a protein that does not intrinsically bind to DNA is not crosslinked to DNA by formaldehyde, even at extremely high (up to 50 mg/ml) protein concentrations. Also, only one intramolecular and no intermolecular methylene bridges were found in native tRNA incubated in 0.2 M formaldehyde at room temperature for 15 days (Axelrod et al. 1969).

The issues delineated here have led us to shift our focus from primary products of formaldehyde reactions with nucleic acids in aqueous solution to the products that may result from conversions during the first step of fixed tissue processing, specifically tissue dehydration in a graded series of alcohols. In this study, we show that “dehydrated” formaldehyde-modified 2′-deoxyadenosine 5′-monophosphate (dAMP) reacts with ethanol, that the reaction is catalyzed by orthophosphate, and that the major product, N⁶-ethoxymethyl-dAMP, is stable under conditions used to reverse formaldehyde modifications of nucleic acids in aqueous solution. In addition to the N,O-mixed acetal, we also found minor amounts of free heterocyclic bases (adenine, N⁶-hydroxymethyladenine, and N⁶-ethoxymethyladenine), as well as cross-linked nucleotides. These findings indicate that histological processing may account for the major alterations seen in nucleic acids recovered from formalin-fixed, paraffin-embedded tissues.

Materials and Methods

Adenine and the sodium salt of dAMP were purchased from Sigma-Aldrich Co. (St Louis, MO). Lithium perchlorate, ethanol (99.5+ %), and acetone (99.9+ %, high-performance liquid chromatography [HPLC] grade) were products of Aldrich Chemical Co. (Milwaukee, WI). Methanol-free 10% formaldehyde, electron microscopy grade, was purchased from Polysciences, Inc. (Warrington, PA). The OLIGOPrep HC cartridge (C₁₈ alkylated porous polystyrene-divinylbenzene copolymer) and chromatographic supplies (HPLC-grade acetonitrile, water, and 2 M triethylammonium acetate, pH 7.4) were purchased from Transgenomic Inc. (Omaha, NE). One molar triethylammonium hydrogen carbonate buffer, pH 8.4–8.6, HPLC grade, was obtained from Fluka Chemie GmbH (Buchs, Switzerland).

A stock solution of unfractionated formaldehyde-modified dAMP (FA-dAMP) was prepared by mixing equal volumes of 1 mM nucleotide in Milli-Q water (Millipore; Bedford, MA) and 7.4% formaldehyde to yield 0.5 mM nucleotide with 3.7% formaldehyde in Milli-Q water. After incubating the solution at room temperature for 3 days, the reaction was quenched by freezing at −20C, at which temperature the reaction mixture was stored.

Aliquots of the FA-dAMP solution, with or without various volumes of admixed 50 mM potassium phosphate buffer, pH 7.0, were treated with 10 volumes of ice-cold 2% LiClO₄ in acetone, kept at −20C for 30 min, and then centrifuged for 15 min at 11,750 X g. After washing the precipitates twice with acetone, some precipitates were kept dry in open vials. Others were dissolved in a volume of 0.1 M triethylammonium acetate (TEAA), pH 7.4, to yield 71.4 μM FA-dAMP, or covered by the same volume of acetone or ethanol. All of the samples were then incubated at room temperature. At the end of the incubation, and immediately before the HPLC analysis, dry samples and precipitates recovered by centrifugation from the organic solvents used were also dissolved in 0.1 M TEAA.

Fractionation of the samples was performed on an OLIGOPrep HC cartridge (7.8 × 50 mm) integrated into a Hitachi D-7000 HPLC System (Hitachi High Technologies America Inc.; San Jose, CA). A discontinuous linear gradient of acetonitrile in 0.1 M TEAA was used: 0–2.5% for the first 5 min, 2.5–7.5% for the next 20 min, and 7.5–17.5% for the last 20 min. An elution rate of 1 ml/min was employed. For the isolation of products formed by FA-dAMP under ethanol in the presence of coprecipitated orthophosphate, 0.1 M TEAA was replaced with 0.1 M triethylammonium bicarbonate. Collected fractions were evaporated under vacuum in a Büchi Rotavapor (Flawil, Switzerland) and then desalted by repetitive evaporation of the residue solutions in Milli-Q water.

Ultraviolet (UV) absorption spectra were recorded on a Beckman DU 600 single-beam spectrophotometer (Beckman Instruments; Fullerton, CA).

Fourier-transform ion cyclotron resonance (FTICR) mass spectrometry (MS) (Comisarow and Marshall 1974; Hendrickson et al. 1999) was performed on desalted fractions that were mixed with 10% isopropanol to provide a final analyte concentration of 10 μM. Analyses were carried out by nano-spray (Wilm and Mann, 1996) on an Apex III FTICR mass spectrometer (Bruker Daltonics; Billerica, MA) equipped with a 7.0 T active shielded superconducting magnet and an Apollo atmospheric pressure ionization source. The homebuilt, heated-metal desolvation capillary was kept at 120–150C. Each experiment was performed in negative ion mode and required loading 5–10 μl of analyte solution into a freshly pulled borosilicate needle, while a platinum wire was inserted from the back end to provide the necessary nanospray voltage. All data were acquired in broadband mode and were processed using Bruker XMASS 6.0.1 software. The tandem MS (MS/MS) experiments were carried out by isolating the precursor ion of interest using correlated radiofrequency sweeps (de Koning et al. 1997) followed by activation through sustained off-resonance irradiation against an argon background to obtain collision-induced dissociation.

Figure 1

Chromatographic profile of FA-dAMP. 1: dAMP; 2: N⁶-hydroxymethyl-dAMP; and 3: N⁶, N⁶-bis(hydroxymethyl)-dAMP. The high-performance liquid chromatography was done using triethylammonium acetate as an ion-pairing agent.

Figure 2

Kinetics of the formaldehyde-freed FA-dAMP demodification in different conditions. Details are given in the text. 1:dAMP; 2:N⁶-hydroxymethyl-dAMP; and 3: N⁶, N⁶-bis(hydroxymethyl)-dAMP.

Results

The stock solution of FA-dAMP used in this study was obtained by a 3-day incubation of 0.5 mM dAMP in unbuffered 3.7% formaldehyde at room temperature. The HPLC profile in Figure 1 demonstrates that the stock was almost entirely composed of N⁶-hydroxymethyl-dAMP and N⁶, N⁶-bis(hydroxymethyl)-dAMP (63% and 37% of the total area under the peaks, respectively), which is in agreement with earlier studies (McGhee and von Hippel 1975). Dehydration of nucleotides in the mixture and removal of unbound formaldehyde were achieved by precipitating the nucleotides out of the stock solution and by washing the precipitate with acetone as described in Materials and Methods. Analyzed immediately after dehydration (profile not shown), FA-dAMP was found to contain dAMP, N⁶-hydroxymethyl-dAMP, and N⁶, N⁶-bis(hydroxymethyl)-dAMP in a 1.4: 71:28 ratio, which suggests partial demodification of the least stable dimethylol adduct.

Next, we used HPLC to analyze the composition of the FA-dAMP precipitates kept dry at ambient humidity (Figure 2), redissolved in 0.1 M TEAA, pH 7.4 (Figure 2), or covered by the same volume of acetone or ethanol (Figures 2C and 2D, respectively). Under all conditions tested, mono- and dimethylol adducts underwent spontaneous demodification, which was fastest in the aqueous solution and slowest in precipitates incubated under the organic solvents. The demodification seen in a sample kept dry (Figure 2) was found to be even further accelerated by coprecipitated orthophosphate as shown in Figure 2E. In that case, 50 mM potassium phosphate buffer, pH 7.0, was added to the FA-dAMP stock solution at a final concentration of 8.3 mM just before precipitation.

When incubation was extended for longer than a week, the FA-dAMP precipitate kept under ethanol accumulated a minor (3.6% of the total area under the peaks) byproduct eluting in 24.2 min (Figure 3). However, when the reaction was repeated in the presence of coprecipitated orthophosphate, the byproduct yield increased nearly 6-fold with just an overnight incubation (Figure 3). Figure 3C shows that this product withstood the 30-min heating at 70C at neutral pH, whereas N⁶-hydroxymethyl-dAMP and N⁶, N⁶-bis(hydroxymethyl)-dAMP present in the initial sample (Figure 3) decomposed.

Figure 3

Chromatographic profiles of FA-dAMP precipitates kept for 8 days under ethanol alone (profile A) or for 20 hr in the presence of coprecipitated orthophosphate (profile B). Profile C corresponds to the latter sample dissolved in 0.1 M triethylammonium acetate, pH 7.4, and then incubated at 70C for 30 min. 1: dAMP; 2: N⁶-hydroxymethyl-dAMP; 3: N⁶, N⁶-bis(hydroxymethyl)-dAMP; 4: a byproduct formed in ethanol.

Figure 4

Effect of neutral phosphate buffer concentration on the composition of the FA-dAMP coprecipitates kept under ethanol for 20.6 hr. 1: dAMP; 2: N⁶-hydroxymethyl-dAMP; 3: N⁶, N⁶-bis(hydroxymethyl)-dAMP; 4: a byproduct formed in ethanol.

Figure 4 and Figure 5 provide further insight into the effects of coprecipitated orthophosphate on FA-dAMP conversions in ethanol. In particular, Figure 4 illustrates the concentration effect of neutral phosphate buffer admixed with the FA-dAMP stock solution before precipitation. The yield of the product with a retention time (t_R) of 24.2 min increased with an increase in buffer concentration, and at 20–25 mM, it reached approximately 40% after a 20.6-hr incubation of the coprecipitates under ethanol. An incubation of 3 days allowed us to detect the slow accumulation of at least 10 additional compounds (see Figure 5). The brief heating of the mixture at 70C and neutral pH resulted in decomposition of mono- and dimethylol adducts and the unknown component with a t_R of 7.29 min (data not shown). This indicated the absence of the heat-labile N-hydroxymethyl group in all the others products. However, when the heat treatment of the same mixture was performed in 40 mM Tris-acetate buffer, pH 4, the subsequent HPLC analysis revealed only two peaks (Figure 5) corresponding to dAMP and a minor component (t_R = 4.95 min) that was present in the mixture before the treatment (Figure 5). This result demonstrated that the majority of the products formed in the FA-dAMP and orthophosphate coprecipitates are acid labile and that the component eluting at 4.95 min appears to be heat and acid resistant. The products marked by their respective t_Rs in Figure 5A were desalted as described in Materials and Methods and then characterized by UV absorbance spectroscopy and electrospray ionization (ESI) FTICR MS (see summary in Figure 6).

A product that was a minor component in the FA-dAMP precipitate kept under ethanol alone but abundant in the presence of orthophosphate was found to have a molecular mass 28 U heavier than that of N⁶-hydroxymethyl-dAMP (Figure 7). This suggests that it was formed by condensation of one molecule of ethanol with either the heterocyclic base or the deoxyribose phosphate of N⁶-hydroxymethyl-dAMP. However, the product fragmentation pattern in Figure 8, provided by tandem mass spectrometry, revealed no anions with the unmodified base, but two anions, 177.00 U and 195.01 U, derived from the unmodified deoxyribose phosphate. Because the product is acid labile (Figure 5) and its UV absorption maximum is at nearly the same wavelength (264.4 nm) as that of N⁶-hydroxymethyl-dAMP (264.8 nm, Rait et al., unpublished data), together the observations are indicative of an N,O-mixed acyclic acetal, specifically, N⁶-ethoxymethyl-dAMP (Bridson et al. 1980; Fraenkel-Conrat and Singer 1988). In agreement with this assignment, structures in Figure 8 specify major anions derived from N⁶-ethoxymethyl-dAMP by collision-induced dissociation (Figure 6, wavy lines).

Figure 5

Chromatographic profiles of FA-dAMP, coprecipitated from 10 mM phosphate buffer, pH 7.0, and then kept under ethanol for 3 days, before (profile A) and after (profile B) a 30-min incubation at 70C in acetate-Tris buffer, pH 4.0. 1: dAMP; 2: N⁶-hydroxymethyl-dAMP; 3: N⁶, N⁶-bis(hydroxymethyl)-dAMP; 4: a major byproduct formed in ethanol; others numbers above the peaks are retention times in minutes. The high-performance liquid chromatography was done using triethylammonium bicarbonate as an ion-pairing agent.

Figure 6

Putative structures and selected characteristics of compounds marked in Figure 5A by retention times. ∗Wavy lines denote bonds whose scission leads to the fragmentation discussed in the text or shown in Figure 8. ∗∗The compound decomposes during desalting of the corresponding chromatographic fractions.

Figure 7

Mass spectrum of N⁶-ethoxymethyl-2′-deoxyadenosine 5′-monophosphate.

Whether both hydroxymethyl groups of the dAMP dimethylol adduct can react with ethanol was determined by analyzing the byproduct with the highest t_R in Figure 5A. First, the molecular mass of the byproduct was found to be 56 U heavier than that of N⁶, N⁶-bis(hydroxymethyl)-dAMP. Second, either of two ethanol residues appeared to be associated with the heterocyclic base. This followed from tandem mass spectrometry that revealed (spectrum not shown) two anions derived, as in the case of N⁶-ethoxymethyl-dAMP, from the unmodified deoxyribose phosphate, and two specific anions of 250.13 U and 160.06 U. The former corresponds to a diethoxymethyladenine anion formed by collision-induced heterolysis of the nucleotide N-glycosidic bond. The latter may represent a similar anion with the exocyclic nitrogen included in the ethylene imine cycle as proposed in Figure 6(wavy lines).

Under the conditions used, the reactivity of N-methylol groups also resulted in cross-linking. According to Feldman (1967) and Chaw et al. (1980), adenine ribo- and deoxyribonucleosides, linked through their exocyclic amino groups, are characterized by UV spectra of a very distinctive shape with maxima positioned near 272 nm. These were characteristics of two byproducts with t_Rs of 27.14 min and 37.71 min in Figure 5A. Based on respective molecular masses of 673.14 U and 731.23 U, the byproducts (shown in Figure 6) originated from reactions of N⁶-hydroxymethyl-dAMP with dAMP and N⁶-ethoxymethyl-dAMP, respectively. It is worth noting that all three nucleotides involved in cross-linking constituted major components of the analyzed reaction mixture (Figure 5). A compound eluting in 33.67 min (0.59% of the total area under the peaks in Figure 5) also had a mass (731.21 U) of N⁶-hydroxymethyl-dAMP cross-linked to N⁶-ethoxymethyl-dAMP. However, its UV absorption maximum was found at 263.8 nm, which indicated a cross-link other than the methylene bridge between two exocyclic amino groups. Clearly, further studies will be necessary to complete the structural characterization of this minor component.

In addition to the products characterized previously, the analyzed mixture was found to contain free heterocyclic bases. The component eluting at 4.95 min (Figure 5) was attributed to adenine because its UV spectrum and t_R coincided with those of the authentic adenine sample. Another component (t_R = 16.74 min) appeared to be N⁶-ethoxymethyladenine, which was supported by an exact mass of 193.10 U and by its transformation into adenine on heat treatment at pH 4. Unlike N⁶-ethoxymethyladenine, the component eluting in 7.29 min was labile and its desalting resulted mostly in demodified adenine. Considering that the detected free bases originated from nucleotides dominant in the mixture, we suggest that this labile component corresponds to N⁶-hydroxymethyladenine. Thus exposure of the FA-dAMP precipitates under ethanol in the presence of orthophosphate led to depurination of the nucleotide and its derivatives.

Discussion

This study was performed on a FA-dAMP preparation obtained in 10% unbuffered formalin and composed almost entirely of N⁶-hydroxymethylated dAMP with, on average, 1.3–1.4 formaldehyde adducts per nucleotide. Although the “fixation” time (3 days) was significantly longer than that in regular histological practice, no cross-linked nucleotides or other products were detected by reverse-phase ion-pair HPLC (Figure 1). When freed from excess formaldehyde and then dissolved in a neutral solution, FA-dAMP underwent demodification (Figure 2), restoring dAMP as expected (McGhee and von Hippel 1975). The slow rate of the reverse reaction observed at room temperature assumes a particular significance for investigations involving the recovery of nucleic acids from fresh formalin-fixed specimens. As shown in Figure 2B, up to 50% of the adenine residues remain as an N⁶-hydroxymethyl derivative after a 24-hr spontaneous demodification. Although higher temperatures have shown to increase the rate of demodification (Haselkorn and Doty 1961; McGhee and von Hippel 1975; see also an example in Figure 3), the extent of this process also depends on the concentration of formaldehyde-modified nucleic acids. For inadequate dilutions, extensively modified nucleic acids could release enough formaldehyde to establish a new equilibrium in this reversible reaction even at high temperatures. Also, to avoid the “poisoning” of enzymes by released formaldehyde, it is reasonable, before PCR or RT-PCR, to precipitate the recovered nucleic acids on completion of their high-temperature treatment.

Figure 8

Fragmentation pattern of N⁶-ethoxymethyl-2′-deoxyadenosine 5′-monophosphate.

In addition to specimen incubation in 10% formalin, the preparation of a tissue for histological study includes three more postfixation steps: (1) a 6- to 24-hr dehydration of the specimen in graded concentrated ethanol, from 70–80% up to 100%; (2) clearing the specimen in a solvent miscible with the embedding medium (e.g., xylene in paraffin embedding); and (3) embedding in melted paraffin at 60C or plastic resin at room temperature (Lillie 1965; Fox et al. 1985). Focusing on step 1, our data show the possible effects of dehydration on formaldehyde-modified adenine residues.

First, the data in Figure 2 demonstrate that dehydration cannot stabilize products formed at the fixation step, and that the rate of demodification in the absence of bulk water could still be as high as it is in aqueous solution. Further, this rate is shown to be highly sensitive to the amount of orthophosphates, (HO)₂PO₂ ¹⁻ and HOPO₃ ²⁻, present as a coprecipitate in the dehydrated FA-dAMP samples (Figure 3). In actual specimens, demodification can be similarly affected by physiological phosphate contained in the tissue to be processed, various phosphomonoesters (e.g., free nucleotides), and by the phosphate buffer (usually 75 mM) used to neutralize the 10% formalin solution.

Second, in FA-pdA samples, dehydrated and kept under ethanol just overnight, we found (Figure 4 and Figure 5) up to 40% N⁶-ethoxymethyl-dAMP, a nucleotide derivative whose existence was never taken into account in the analysis of nucleic acids recovered from formalin-fixed specimens (Douglas and Rogers 1998; Srinivasan et al. 2002). Meanwhile, because of their bulkier size, N-ethoxymethyl groups can degrade the nucleic acid matrix properties more efficiently than N-hydroxymethyl groups. Furthermore, N-ethoxymethyl groups are remarkably stable in neutral and alkaline media. For example, the half-times of hydrolysis of N⁶-ethoxymethyl-2′,3′-O-isopropylidenadenosine were found to be 10 hr and 1.5 min at 20C in 50 mM NaOH and 50 mM HCl, respectively (Bridson et al. 1980). Yet, even in acidic media, demodification of analogous derivatives of cytosine and guanine nucleosides could be troublesome because of their, respectively, 80- and 13-fold longer half-times of hydrolysis (Bridson et al. 1980). Because no such products were reported in previous studies using water/ethanol mixtures (Fraenkel-Conrat and Singer 1988), it is plausible that formation of the N-ethoxymethyl adducts may only occur toward the end of specimen dehydration (and at the beginning of its rehydration)—i.e., under anhydrous ethanol conditions. Hence, it is reasonable to suggest that tissue dehydration results in molecular dehydration by transforming N-hydroxymethyl groups, -NHCH₂OH, into Schiff bases, -N = CH₂. In such a scheme, the bulk anhydrous alcohol acts as a medium to effectively absorb the water of the molecular dehydration and, additionally, as an abundant reactant to compete with a nucleotide exocyclic amino group for the acid-activated Schiff-base, -HN⁺ = CH₂↔-NH-CH₂ ⁺ (Wagner 1954).

Third, the character of the products that are formed in addition to N⁶-ethoxymethyl-dAMP (Figure 5A and Figure 6) led us to conclude that dehydration can also be responsible for nucleic acid degradation and cross-linking. The detection of adenine, as well as N⁶-hydroxymethyladenine and N⁶-ethoxymethyladenine (3.1% of the total peak areas in Figure 5), is a clear sign that the nucleotide and all its derivatives in the FA-dAMP preparation undergo a rather nonselective depurination when kept under ethanol. Although the content of the detected free heterocyclic bases is comparatively small, even a 1% depurination in the case of polymeric DNA could produce fragments of no more than 100 nucleotides, on average.

Cross-linking also appears to be nonselective because N⁶-hydroxymethyl-dAMP reacted with either of two other major components of the incubated FA-dAMP—i.e., with the unmodified nucleotide and its N⁶-ethoxymethyl derivative. Because the formaldehyde-induced cross-linking of nucleotides has long been considered as an irreversible or, at least, as extremely stable modification (Collins and Guild 1968), the ability of the pH 4 treatment at 70C to demodify the methylene bis-nucleotide adducts in 30 min is surprising. It seems that the hydrolytic stability of the -NH-CH₂-NH- group, which was previously tested only with cross-linked nucleosides in 0.5–4 N acids (Feldman 1962; Chaw et al. 1980), is closer to that of the -NH-CH₂-O- group than is generally thought. Further, the -N(6) H-CH₂-O- group and the nucleotide intrinsic fragment N(9)-C (1′)H-O- both qualify as N,O-mixed acetals (acyclic and semicyclic, respectively). As such, their formation and hydrolysis are often subject to catalysis by a Brønsted acid or a Lewis acid (Gabbutt and Hepworth 1995; Sugiura et al. 2001). Therefore, acid salts of orthophosphoric acid (probably in the form of crystalline hydrates) and/or its monoesters could play this catalytic role in all the conversions reported here.

It is worth remembering that phosphate-buffered formalin was introduced in tissue fixation with the aim of neutralizing the formic acid contaminating formalin long before the first PCR and RT-PCR trials on nucleic acids recovered from formalin-fixed, paraffin-embedded specimens (Impraim et al. 1987; von Weizsacker et al. 1991). According to Lillie (1965), phosphate buffering “…resulted in a definite increase in frequency of demonstrability of ferric iron in blood pigments, and it almost entirely prevented formation of the so-called formalin pigment.” However, Fox et al. (1985) commented that prevention of “formalin pigment” formation was essential only when dealing with blood-rich tissues.

As it becomes clear that the presence of endogenous or exogenous acids during dehydration promotes ethoxylation, cross-linking, and depurination of formaldehyde-modified nucleic acids, the urgency of finding alternative reagents or changing the established procedure becomes more pressing. It is likely that the body of empirical observations accumulated to date in formalin fixation might already contain various solutions for this problem. Watson et al. (1986) suggested abandoning the entire postfixation processing in favor of the long-term storage of fixed brain tissue at −20C in an ethylene glycol-based cryoprotectant solution. It was confirmed later (Lu and Haber 1992; Hoffman and Le 2004) that this laborsaving approach did not compromise mRNA levels detected by in situ hybridization after 3, 10, and even up to 20 years of storage. Basyuk et al. (2000) increased the sensitivity of in situ hybridization with RNA probes 5- to 6-fold by employing formalin fixation in phosphate buffered saline, pH 9.5. Potentially, alkaline fixation might reduce or even abolish unwanted conversions of N-hydroxymethyl groups during conventional processing. Last, Fang et al. (2002) have recently reported that the quality of extracted DNA could be dramatically improved if dehydration was started from 30% ethanol, continued in 10% steps, and completed by critical point drying. This course of action is more likely to lead to the elimination of low molecular weight acids during the early dehydration steps, thus excluding their coprecipitation with nucleic acids that could occur in regular 70% or 80% ethanol. In addition, critical point drying appears to accelerate the decomposition of N-hydroxymethyl groups by removing the spontaneously released formaldehyde from specimens.

In this study, we have identified specific nucleotide alterations that are not reversible by methods, including enzymatic and heat treatment methods, currently used in the isolation of nucleic acids and their preparation for molecular biological analysis. It is possible that these modifications, were they to occur in tissue, would impair the ability of transcriptases to make cDNA copies of endogenous ribonucleic acids. Even if only 1% or 2% of bases are so modified, the transcription and amplification of sequences more than a few hundred bases long could be significantly impaired. It is plausible, therefore, that the modifications described here could explain the difficulty in recovering long transcripts, or even detecting low-abundance RNA species, from formalin-fixed, paraffin-embedded tissues. Additional experiments on polynucleotides will be required to establish whether these or similar changes actually occur during tissue processing.

In conclusion, we express a hope that extensive mechanistic studies on the effects of alcoholic dehydration on conversions of formaldehyde-modified nucleic acids can fill the gap in our understanding of the status of nucleic acids in archived formalin-fixed, paraffin-embedded specimens. This would greatly improve the quality of molecular biological analysis in retrospective studies on such specimens and would also result in a tissue fixation procedure that satisfies the needs of tissue morphologists and molecular biologists alike. Additional experiments on oligonucleotides and intact nucleic acids will be required to determine with certainty the extent to which the modifications occurring during dehydration impair molecular biological analysis of formalin-fixed, paraffin-embedded tissue.

Footnotes

Acknowledgements

This work was supported, in part, by grants from the National Cancer Institute (1R33 CA107844 and 1R21CA91227) to T.J.O. and by the American Registry of Pathology.

This work is not to be construed as official or as representing the views of the Department of the Army or the Department of Veterans Affairs.

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